Measure the length of 1mm using a ruler or caliper. In this case, 1mm is magnified to 108.5mm.

Then measure the size of the structure on paper. In this case, we look at stomates from the bottom of a leaf. The guard cells are 3.6mm long.

• Place a ruler on the stage and take a picture. A full unit (1 mm) should be visible. Transparent ruler are better, otherwise it’s not possible to see the markings. Take a digital photograph of the ruler.
• Print the micrograph of the ruler.
• Take a picture of the specimen. Make sure that you use the same magnification.
• Print the picture of the specimen and be sure that the size of the picture is the same as the size of the picture of the ruler. Do not change the size of the print out.
• Now it’s time for a little math. Use the ruler and measure out the size of the 1mm on the print out. Measure it out in mm. Let’s call this “r”.
• Measure out the size of the structure that you want to determine. This is “s”. Make sure that you use the same units (mm)!
• The real size of the structure in mm can be calculated as follows: size = (1mm * s) / r

Let’s use the example in the pictures on the left:
size = (1mm * 3.6mm) / 108.5mm
size = 0.03mm = 30 micrometers

Samples that are prone to form air bubbles

Not all specimens are the same. Some specimens can be the cause for more air bubbles than others. This depends on a variety of factors. The following characteristics may result in more bubbles:

• Large sheet-like specimens (e.g. onion skin): These specimens may catch air bubbles underneath them and prevent them from escaping. Push out the air bubbles before adding a cover slip.
• Specimens with many fine hair: The hair catch much air and prevent the water from reaching all the parts of the specimen. The surface tension of the water is too high, and the water therefore does not “flow” into all parts of the specimen. This is comparable to the “Lotus Effect”, where the water does not wet the surface of the lotus leaf.
• Fatty and hydrophobic specimens: These too do not accept water well, especially if the surface area of the specimen is large (many fine hair, etc). It may help to treat the specimen in alcohol or an alcohol-water mixture to remove the fatty surface.
• Porous specimens: The pores of the specimen may be filled with air, which can be difficult to remove. The cells of plant stems, the vascular tissue, for example, are able to hold air. It is possible to remove the air by placing the specimen into a vacuum while it is submerged in the fixing solution. Aspirators (eductor-jet pumps) can be mounted to a water tap to produce a vacuum.

Why air bubbles should generally be avoided

Some air bubbles are certainly tolerable and unless one wants to produce high-quality pictures it is often not worth the effort to make a completely bubble-free specimen. It is easily possible to simply move the slide and observe a different part of the specimen. Generally, air bubbles should be avoided, especially by beginning microscopists, who may have a problem distinguishing bubbles from the real specimen. The reasons why air bubbles can be problematic are:

• Bubbles hinder the free movement of organisms, such as ciliates
• The bubbles cause optical artifacts at the place where the air meets the water. The air bubble appears to be surrounded by a dark ring. This dark ring covers some parts of the specimen and makes observation more difficult.
• The microscope optics are designed to give optimum resolution for a specimen which is surrounded by water. If the bubble is large and the specimen completely surrounded by air, then the resolution is lower.

Are there cases when air bubbles are beneficial?

Under some rare circumstances, air bubbles can even be beneficial. The bubbles can serve as a source of oxygen for some organisms, such as paramecia and other ciliates. It is possible to see them collect around the bubbles. Air bubbles are also easily viewable and can therefore help beginners to more easily find the correct focus. Naturally, the bubbles should not be confused with the actual specimen, something that beginners sometimes do because the bubbles are so prominent and can be seen even if the specimen itself is not in focus.

How to minimize air bubbles in wet mounts

Needless to say, the preferred method depends on the characteristics of the specimen. Try out the following:

• Cover slip placement: Lower the cover slip on the water droplet with an angle. This permits air to escape on one side.
• Water placement: If the specimen is not fully submerged in the water droplet, add another droplet on top of the specimen before lowering the cover slip.
• Immersion oil: Use a medium other than water. Try immersion oil, which is hydrophobic. Some specimens prefer water, others oil.
• Break the surface tension: Add a small amount of detergent, such as soap. This will break the surface tension of the water. The water will therefore adhere better to some specimens, thus preventing bubbles. The soap may also harm some water organisms, however.
• Apply a vacuum: This speeds up the movement of the fixing solution or water into the specimen.
• Dehydrate the specimen: Place the specimen into alcohol. Some specimens will shrink and lose water and air. By placing the specimen into water again, the specimen will take up the water.
• Remove oil and fat: Wash the specimen in alcohol.
• Add water: If the air bubble is large and reaches the side of the cover glass, you can add more water from the side of the cover glass.

A few days ago I ordered a microtome. Here is a video showing you the different parts: Parts of a Microtome

Now it’s time to test the device. The first sample is a carrot. It can be cut into the right shape to fit into the specimen holder of the microtome and it is sufficiently solid to allow for easy cutting, but not too hard. Carrots can also be used to hold other specimens. In this case the “carrot cylinder” is cut in half and the specimen can be inserted between the carrot halves. The carrot acts as a support.

A hand microtome (or cylinder microtome) is a device used to make thin cuts of a specimen for microscopic observations. In the video I am unpacking a new hand microtome and showing the different parts:

• The clamp: This one is optional, but very useful. It holds the microtome to a table. It adds stability and convenience, thereby making the microtome cuts more reproducible.
• The knife: This one looks like an old fashioned razor knife.
• The microtome: It has a central hole into which to place the specimen. A screw at the opposite end moves a piston up, which in turn pushes the specimen up. The plate of the microtome acts as a guide for the knife.
• The mold: A small brass cylinder serves as a mold for making paraffin blocks containing the specimen. This paraffin block is then inserted into the hole of the microtome.

What is a wet mount?

In a wet mount, the specimen is suspended in a drop of liquid (usually water) located between slide and cover glass. The water refractive index of the water improves the image quality and also supports the specimen. In contrast to permanently mounted slides, wet mounts can not be stored over extended time periods, as the water evaporates. For this reason, a wet mount is sometimes also referred to as a “temporary mount” to contrast it from the “permanent mounts”, which can be stored over longer times. The permanently mounted slides use a solidifying mounting medium, which holds the cover glass in place. The naming can be a bit problematic, because it is also possible to make wet mounts that can store over extended time periods. These are special cases, however.

Different types of wet mounts

Wet mounts can be made using several different kinds of liquids. Water, immersion oil and glycerin (glycerol) can be used, with water probably being the most commonly used. The source of the water is quite important, especially when observing living specimens. If you use water with a wrong osmotic potential (ie. too much or too little salt and mineral content), then there is the danger of damaging the specimen. A too high salt content can result in the specimen to lose too much water. Too low a salt content, and the specimen may swell and burst.

• Using water from the natural habitat of the organism: In the case of water organisms, such as algae or ciliates, the liquid water should come directly from the sample. In this case the organism is immersed in its own natural environment. The microscopist uses a dropper to place a drop of pond water directly on the microscope slide.
• Using 0.9% salt water: In some cases water from the natural habitat may not be available. This is the case when observing bacteria or molds grown on petri-dishes. Yoghurt bacteria, for example, need to be diluted a lot before being able to observe them, otherwise they are too dense to be observed as single cells. In this case it is necessary to mix some salt (NaCl) into some water to ensure an optimal osmotic potential. This “physiological saline”, as it is called, can be made by dissolving 9 grams of table salt (NaCl) in 1 liter of water (or 0.9g Nacl in 100ml of water).
• Using tap water: If one wants to observe non-living specimens, such as dust samples, sand grains, or thin section cuts of plant material, then it is also possible to use regular tap water. These specimens are not osmotically sensitive. If the specimen is observed without water, in a dry condition, then the resolution and image quality may not be sufficiently high. I advise you to try out both to see the difference. The following post includes images of pollen grains mounted in air and water, for comparison: The effect of the mounting medium on specimen and image quality
• Using immersion oil: Some wet mounts are not made with water, but by using immersion oil. Immersion oil is usually placed on top of the cover glass. In this case the specimen does not get into contact with the oil. It is also possible to submerge the specimen in the oil, however. Heat-fixed bacteria can be observed directly by placing a drop of immersion oil on the specimen, without cover glass. The oil-immersion objective is then rotated directly into the oil for observation. It goes without saying, that this procedure can only be used for specimens that do not contain water (and are, therefore, not living). It also only works for specimens that stick to the glass slide – there is no cover glas. If you need to observe these specimens with a lower magnification (ie. no immersion objective), then one needs to use a cover glass, of course. Other specimens, such as synthetic textile fibers, are hydrophobic in nature, and do not like to be mixed with water. They tend to float on top of the water drop and this can be cause for air bubbles. In this case I also recommend to use immersion oil and a cover glass to keep the sample flat.
• Pure glycerin or glycerin-water mixtures: Glycerin has a strong tendency to withdraw water from the sample. For this reason it also acts as a preservative. On the down side, the glycerin may therefore cause the specimen to shrink and deform. Especially algae and other water organisms are sensitive to dehydration. Other specimens, such as sectioned or microtomed plant material are not as sensitive. The reason why glycerin is used is because of its high refractive index. This may be necessary to see certain structures. If a lower refractive index is needed, then one should mix some water into the glycerin. It is possible to seal the glycerin mount by applying nail polish to the sides of the cover glass. This will hold the cover glass in place for longer time periods. This is then an example of a wet mount, which was made into a permanent mount.

Advantages and disadvantage of a wet mount

Compared to permanently mounted slides, wet mounts do have certain advantages:

• Quick preparation: specimen fixation, dehydration and staining are not necessary (but possible, if required). For this reason, wet mounts are the first kind of mounts that students learn to make.
• Few artifacts: If there is no chemical and physical processing of the specimens before observation (no fixation), there are little artifacts and the specimens appear in their natural condition.
• Living and moving: It is possible to observe living and moving organisms. It is also possible to observe certain processes of life, such as feeding, cell division etc. (for water-based mounts)
• Natural colors: The colors are natural and not faded. The colors of permanently mounted specimens may fade over time.

Disadvantages of wet mounts include:

• Movement: The advantage of observing movement can also be a disadvantage. Due to the movement of the organisms it may be more difficult to take pictures or to make drawings. There is a solution to this problem: one can slow down ciliates and other protozoa by adding a solution such as ProtoSlo, which increases the viscosity of the water.
• Evaporation: The heat of the lamp causes the water to evaporate more quickly. More water must be added under the cover glass from time to time.
• Focus: Some organisms may swim vertically in the water and therefore move in and out of focus. Here it is important not to use too much or too little water. Too little water may squeeze the specimen between cover glass and slide.
• Storage: Wet mounts can not be stored over a longer time.

Materials and Method

For making a wet mount you need these materials:

• Microscope slides
• Cover glasses
• The specimen to be observed: make sure that the specimen is sufficiently small and thin. Thick specimens must either be cut (microtomed) into sections, be squeezed or torn apart.
• Water: take care that the osmotic potential of the water is compatible with the specimen. For example, do not use fresh water with marine specimens, and vice versa. Use pond water (and not tap water) for observing pond organisms.
• Droppers, pipette: these are for transferring the water
• Tweezers: for handling the specimen, the cover glass and for adding water

If the specimen is already in water (algae, ciliates etc.) then you can proceed the following way:

1. Place a small drop of sample fluid (containing the specimen) in the center of the microscope slide.
2. Hold the cover glass on one side with the help of tweezers. Lower the cover glass onto the water drop at an angle.
3. Then slowly lower the cover glass into the liquid. This will minimize disturbing air bubbles.
4. Remove excess water with filter paper or tissue paper. The cover glass should not float freely. The surface tension of the water should hold it in place. Alternatively you can add more water using a pipette or tweezers.

If the specimen is not in water:

1. Place a small drop of water (without specimen) in the center of the microscope slide.
2. Place the specimen into the water.
3. Add some more water on top of the specimen and make sure that the specimen is completely submerged. Otherwise there is the possibility for air bubbles forming between cover glass and specimen. The remaining steps are the same as above.
4. Hold the cover glass on one side with the help of tweezers. Lower the cover glass onto the water drop at an angle.
5. Then slowly lower the cover glass into the liquid. This will minimize disturbing air bubbles.
6. Remove excess water with filter paper or tissue paper. The cover glass should not float freely. The surface tension of the water should hold it in place. Alternatively you can add more water using a pipette or tweezers.

If you are using a dry specimen (dust, insect parts, etc.), then place a small drop of tap water

How to prevent drying out

The heat of the microscope light will evaporate the water relatively quickly. There are several possibilities to counteract this:

• Keep adding more water from the side of the cover glass. Surface tension will pull the water in.
• Seal the sides of the cover glass with a thick layer of Vaseline (petroleum jelly). Press the cover glass against the slide so that the vaseline is able to seal off the water from the outside.
• Use nail polish to seal off the cover glass. This is used when making wet mounts with glycerin. Keep the glycerin drop very small. The nail polish will not stick to those parts of the cover glass and slide which came into contact with the glycerin.
• Use slides that have an indentation (concave) and are therefore able to hold more fluid. This only works for some samples because the liquid layer may be to thick. These slides are more expensive.
• Use two additional cover glasses to support a third cover glass left and right. These two cover glasses serve as a distance holder for the third cover glass. This way the third cover glass does not float freely on the liquid but is held in place by the two supporting glasses. More fluid can be stored in a stable manner.

Characteristics of a chemical fixative

A good fixing agent should fulfill several criteria:

• It must kill the specimen quickly: But be careful, some chemical fixing agents are toxic and are also harmful to the health of a person.
• It must preserve the structures of the specimen, without introducing deformations or other artifacts. Insects may pull together their appendages, making them more difficult to see. The structures should then be sufficiently stable to withstand the dehydration and mounting.
• It must enter the specimen well to react with all parts: This can be problematic with some specimens. Make sure that the specimen is sufficiently small. Alternatively it is possible to puncture the specimen (insects) so that the fixing agent can enter more easily. Some specimens may contain air bubbles which prevent the fixing agent to reach all parts. In this case it may be necessary to apply a vacuum to remove the air.

Types of fixing agents

Chemical fixing agents can be categorized into the following 4 groups:

• Alcohol and acetic acid: This combination denatures proteins. The alcohol also removes some lipids. This is probably the preferred fixing agent for hobbyists, because it is less toxic than some other fixatives.
• Aldehydes (such as formaldehyde – toxic!): these react with amino groups in the specimen.
• Oxidation agents: these react with lipids.
• Tanning agents: react with proteins and with amino groups.

The choice of the fixing agent must be carefully matched with the specimen. Some fixing agents (eg. alcohol) may result in the shrinking of the specimen and therefore introduces artifacts. Sometimes it may be necessary to gradually increase the concentration of the fixing agent in order to prevent the formation of artifacts, but this depends much on the type of specimen used. I can not give general advice here, and recommend that one consults specific laboratory manuals.

Using alcohol

For the hobbyist who wants to prepare a slide every now and then, keeping a whole set of different chemical fixatives is probably an overkill (and not healthy either). I keep a small bottle of 96% rubbing alcohol on my shelf, into which I drop the specimens, usually small insects, as they arrive. They will store nearly indefinitely in this solution. When For making permanent slides, I directly transfer them into Euparal mounting medium.

Pure alcohol (ethanol) is also suitable for fixing and storing plant specimens, without cell contents. The alcohol has the tendency to shrink the cytoplasm, but does not affect the cell walls. The alcohol also hardens the plant material, making it easier to cut with a microtome (which often removes the cell contents anyway).

Alcohol/acetic acid solution

Acetic acid (acetate) compensates the shrinking effect of the alcohol. The Carnoy Clarke solution uses 3 parts 92% rubbing alcohol mixed with one part pure acetic acid. The correct alcohol:acetate ratio should be fine-tuned experimentally. If the cytoplasm still shrinks too much, the recipe according to Farmer may be tried out (2:1 alcohol:acetate ratio). Fixing should take place for about 24 hours.

After fixing

There are two more steps necessary: the fixing agent has to be removed (washing) and the specimen has to be dehydrated. Several fixing agents are water-based and this water has be be removed before mounting them in a non-water based mounting medium. Dehydration is not necessary when mounting in a water-based mounting medium such as glycerin gelatin. Dehydration is commonly done by placing the specimen in successively higher concentrations of ethanol. Afterwards the specimen is transferred into a solvent which is compatible to the mounting medium. Some mounting media require the specimen to be submerged in xylene (toxic). Other mounting media are able to directly accept the specimen from the alcohol (Euparal). If one sees a clouding of the slide, then this can be an indication that there was still some water in the specimen.

Heat-fixing of bacteria

Bacteria are treated differently. They must not only be killed, but also physically fixed to the glass slide. Otherwise they will be washed off during the staining process. This method also works with cells collected from the inside of the cheek and water samples.

• Place a bacterial suspension on the slide and let dry. Dry gently, dry completely but do not heat, otherwise the cells may pop open.
• Pull the glass slide through the flame of a Bunsen burner (1-2 times). The specimen should not come into contact with the flame (specimen on top, flame on the bottom). This step is called “heat fixing”. It kills of the bacteria and binds them to the glass slide much like an egg to a frying pan. The glass slide should be so hot that you are just able to hold it in the palm of your hands without causing burns. Heat the slide too much and you end up burning the bacteria 8and destroying their structure).
• The bacteria can now be stained. Place a drop of the staining solution on the cold slide. Rinse off with water and dry it in air. Do not dry-wipe, you will remove the fixed bacteria. You can then observe the bacteria directly in oil immersion even without a cover glass. Place the immersion oil directly on the fixed and stained bacteria.

The general procedure of making a wet mount

1. Place a drop of water on the center of the slide. It is also possible to first place the specimen on the slide, but small specimens usually separate more easily from the tweezers or needle if dipped into the drop of water.
2. Place the specimen into the drop of water and if the specimen floats, add another drop of water on top of it. This reduces the possibilities of air bubbles forming.
3. Carefully lower the cover glass so that it touches with one side the drop of water. The cover slip should form an angle of about 45 degrees with the slide. Touch the cover glass on the sides only to prevent finger prints. Alternatively, use tweezers to hold the cover glass.
4. Then lower the cover slip completely. Placing the cover slip at an angle prevents the formation of air-bubbles.
5. Remove excess water with a filter paper or tissue paper

Possible problems of making a wet mount

• The cover glass floats and moves: This is due to too much water. Remove water with the help of a tissue paper. Under no circumstances should there be water droplets on top of the cover glass. This water may get into contact with the objectives.
• The liquid streams and does not settle: This could be due to evaporation. Add more water between coverslip and slide.
• Air bubbles start to become visible: If bubbles were not present before and start to form, then this could be an indication of oxygen production due to photosynthesis. This depends on the oxygen saturation of the water and the amount of photosynthetic algae present.
• Air bubbles are present: Often the cover glass was not lowered from the side at an angle, but placed horizontally on the water drop. It may also be that the the specimen is hydrophobic (fatty) and /or fluffy. In this case, the the water may have problems reaching all of the areas of the speciemen and there is much air caught by the fine structures. Wet the specimen briefly in alcohol and then transfer directly from the alcohol to water. Alternatively you can try to break the surface tension of the water by adding a small amount of surfactant, such as soap or shampoo. Be aware that alcohol or soap may have adverse effects on living organisms.

There are several methods of preparing pollen grains, each one offers advantages and disadvantages. I can not give a general rule, it simply depends on the goal of the investigation and on the sample investigated. Pollen from wind-pollinated plants taken from a dry environment are probably best left in a dry condition, and not mixed with a water-based mountant, which may cause them to swell (depends on the osmotic potential of the medium, however). On the other hand, the obtained image quality and resolution may not be satisfactory in such a dry mount. It is a compromise, in which several factors have to be taken into consideration. A microscopy enthusiast, who does not need the slides for identification purposes, will again set different standards (such as avoidance of toxic solvents). People who want to publish their results, in turn, may have to rely on the preparatory technique which is customary in their field of research, for reasons of comparison. I recommend that the different methods are tried out.

Mounting techniques

Glycerol wet mount: Place a small drop of glycerol on a clean slide and tap the anthers of the plant so that the pollen falls into the glycerol. If necessary, carefully separate large chunks of pollen grains by stirring. Place a cover slip on top and seal the sides of the cover slip with nail polish. Use a very small amount of glycerol to make sure that the nail polish has enough area to stick the coverslip to the slide. Glycerol wet mounted slides can be stored for months if there is no leakage. The glycerol will withdraw water from the pollen. If the pollen is not dry, then there is a possibility of the pollen to shrink.

Air mounts (dry mounts): In this case, no liquid mounting medium is used. A cover slip is placed on top of the pollen grains and sealed on the side, either with nail polish or with tape. Nail polish may flow very quickly between cover slip and slide, so it may be best to use a nail polish of low viscosity (by letting some solvent evaporate first).

Glycerol jelly (according to Kisser): This is a very popular mounting medium for pollen. It is phenol-free (antiseptic additive) and therefore non-hazardous. It contains 10g of gelatin, 35ml distilled water and 30ml of glycerol (glycerin). After mounting, the sides of the cover slip need to be sealed. Due to the lack of an antiseptic, it is also necessary to work in a sterile manner, otherwise there is the risk of fungal growth in the medium. Maybe it is a good idea to treat the pollen grains first in alcohol to reduce the chance of fungal contamination by spores. Alternatively, one could experiment by increasing the concentration of glycerol.

Non-water-based mounting media: Euparal is a mounting medium which is not water based. Specimens which are present in alcohol can be directly transferred to Euparal. Place a pollen suspension on the slide and let the alcohol evaporate. Before mounting pollen in Euparal, I recommend that the pollen are first washed in alcohol and then compared to the original shape. Does washing in alcohol result in an unacceptable shrinking of the pollen or unacceptable loss of pigments? If not, then mounting the pollen in Euparal may be an alternative.

Reading materal

I found the following article: Marvels of pollen shown by your microscope (Popular Science, September 1939)
(The article recommends the use of organic solvents (such as xylol/xylene and others) to remove oil from the pollen. I do not recommend this due to health reasons, especially when preparing samples for educational purposes. Still, it gives a nice overview of the topic.)

Toxicity: Solvent-based mounting media (such as Eukitt and Canada Balsam) require the specimen to be in xylene prior to embedding. This substance is toxic. Other mounting media, such as Glycerol jelly, may contain hazardous antiseptics. This aspect of toxicity is something to consider when making permanent mounts either as a hobby or for educational purposes in schools. One should ask oneself, if one should not use other alternatives.

Refractive index: The correct refractive index (RI) of the mounting medium can be critical for seeing details of the structure. If one uses phase contrast microscopy, then the RI of the mounting medium should be very different from the RI of the specimen. For regular bright-field work with pigmented specimens, the RI should be the same. In an ideal world, the mounting medium should be matched with the type of specimen. For amateur or educational work, this may be of less relevancy, however. Some high-end microscope objectives are calibrated to be used for a specific RI of the mounting medium, otherwise the resolution is reduced.

Compatibility with specimen: Specimes which are kept in water should be transferred into a water-based mounting medium. Transferring them into a solvent-based mounting medium may result in a clouding of the resin. Likewise, specimens which are kept in alcohol should be transferred to xylene and then embedded in a solvent-containing mounting medium. Euparal allows the specimen to be present in alcohol.

Pigment stability: Some mounting media cause a fading of pigments and stains over time. If pigment stability is of relevancy, then one should use mounting media which do not react with the pigments of the specimen. In some cases a fading of pigments is desirable, however. This brightens the specimen and makes it more easy to observe.

Shrinkage: Some mounting media shrink when they dry. The effect is particularly noticeable when thick specimens (e.g. whole insects) are embedded. Non-water based mounting media are known to do this. Glycerol jelly, which is water-based, does not shrink, however.

Durability: How long should the permanent slides be stored? Non-solidifying mounting media may not hold the specimen in place very well and there is the risk of running out if not sealed properly. Other mounting media may start to crystallize over the years. Still others may adversely react with the pigments of the specimens. Canada balsam is known for its good durability.

Cost: Some mounting media (such as Canada Balsam) are quite expensive. Others can be made in the kitchen from readily available materials (Glycerol jelly).

Ease of use: Here we have to consider two aspects, the preparation of the specimen prior to mounting and the actual mounting process. Some mounting media require the specimens to be dehydrated and fixed before mounting (for resin-based media). This can be a time consuming process. During the mounting process, some media are more prone to form air bubbles (Glycerol jelly).

• Observing dust samples: Students should collect house-dust and bring it to class to be observed under the stereo or compound microscope. Careful, some people may be allergic to dust!
• Observing sand and soil samples: Students should collect sand and soil samples to be observed under the stereo microscope.
• Observing textile fibers: Observing various fibers obtained from clothing (cotton, polyester, nylon etc.). Different colors and textures become visible under the microscope.
• Which printer is the best? Students bring in print-outs of different pictures on different types of paper. The printing resolution can be observed under the stereo microscope.
• Observing water life: A large jar is filled with pond water and a little soil. Algae and other organisms will (hopefully) develop over the course of a few weeks. Do not let the water rot!
• Fungi from cheese: Camembert, Brie, etc. contain edible molds (not hazardous) and can be used. Much safer than rotting food and observing the molds.
• Vegetables and fruits: The teacher cuts the tomatoes and mushrooms in various ways, they can be observed under the stereo microscope. Do not eat the food afterward, you never know what chemicals were left behind on the microscope by previous classes…..
• Hair samples: Each student donates one hair and then they have to match them with the hair left behind on the “crime site”. This is a playful approach into forensics and gives the observation some purpose. Maybe a competition between different groups is also a nice idea. The teacher may have to prepare a set of permanent slides with some hair samples.
• Coins: Coins collect many scratches (and dirt) over the years. How can the scratches be quantified? Is it possible to predict the age of a coin by looking at the number of scratches? The year is imprinted in the coin.
• Observing human cheek cells: This is a classic, really. Using a cotton swab, some epithelium cells from the inside of the mouth are collected and transferred to a microscopic slide.

Things NOT to observe – Some specimens or samples should not be observed in a classroom setting:

• Spoiled food material: they contain hazardous bacteria and fungi. Spores are unhealthy to breath in.
• Body parts: Samples taken from wounds (pus etc).
• Blood samples or other body fluids.
• Urine: Some students (often boys…) may be interested in observing their own urine. Fresh urine should be free of microorganisms (unless there is an infection) and it is not an interesting sample to be observed.
• Animal wastes: Excrements of animals are prone to contain parasites and are a clear health hazard.
• Polluted water Water from polluted rivers, lakes may contain toxic substances and harmful microorganisms. Leave stuff like this to university-level students, who (should) know appropriate safety procedures.

A variety of different standards exist:

• Standard slide: 26 x 76 mm
• Geological slide: 75 x 50 mm
• Petrographic slide: 46 x 27 mm
• Thin sections slide: 48 x 28 mm

Microscope glass slides may be modified in a variety of ways:

• They may have a central indentation to carry several drops of liquid.
• They may have a frosted side to allow for easier writing with a marker.
• They may have polished corners to reduce the possibility of injury due to sharp corners.

Cover glasses (cover slips) exist in a wide range of different sizes, square, round, rectangular. Common sizes include:

• 18x18mm
• 20x20mm
• 22x22mm
• 24x24mm
• various rectangular sizes up to 24x60mm to cover nearly the whole slide.

Choose a cover glass that corresponds to the size of the specimen and the slide. The thickness of the cover glass is important, as it has a significant impact on the resolution of the image. The thickness should correspond to the thickness indicated on the objective lens. In many cases, the cover glass is 0.17mm thick, but there is often a small variation even in the same batch. For critical purposes, it may be necessary to measure the thickness of the individual cover glasses to find one close to the desired thickness (use a vernier caliper to determine the thickness).

Materials: A large glass jar, fresh and unfertilized garden soil, water, hot plate, celophane foil

Method:

• Fill the glass jar with a few centimeters of the garden soil.
• Add non-chlorinated tap water to the soil and fill the jar with the water (3/4 full).
• Boil the soil-water mixture for about 30 min. This will kill off bacteria in the soil and will extract nutrients from the soil. Bacterial spores may survive the boiling, as they are heat-resistant. This is not a problem, though. These bacteria will serve as a food for other microorganisms later on.
• Cool the water to room temperature and let the soil settle to the bottom of the glass jar. Do not filter the soil away. The soil will continue to supply nutrients and will act as a buffer.
• Add a small amount of pond water which contains algae. Do not add too many algae. You may want to scrape off some algae from rocks or take a few algal filaments floating in a pond.
• Cover the jar with celophane foil. This will allow for gas exchange and prevent dirt and dust falling into the water. It also reduces evaporation.
• Wait a few weeks for the algae and ciliates to develop. With a bit of luck, paramecia will grow and form white clouds in the water. The color of the water may also change, an indicator for algal growth.
• Store the jar in a bright place but not in direct sunlight.
• Using a pipette, extract some of the microorganisms to be observed under the microscope.

Troubleshooting:

• Microorganisms do not form: This is probably due to the fact that there were none or not enough in the pond water which was added.
• The water starts to smell bad: This may be due to the system becoming anaerobic. Make sure that enough oxygen is able to enter the water. Paramecia and other ciliates are probably dead by now…..

A specimen for compound microscopy must fulfill several criteria:

• It must be sufficiently thin.
• It should not be too dark (too heavily pigmented).
• If it is not pigmented at all, then it should possess a different refractive index compared to its surrounding medium, otherwise the structure is invisible.
• It should possess sufficient color contrast.

What should one do if the specimens do not fulfill the above criteria? It depends on the type of specimen.

• Thin and strongly pigmented specimen: bleaching. Depending on the type of specimen, different bleaching methods can be used. It is also possible to remove some pigments (such as chlorophyll of plants) by immersing the specimen in alcohol.
• Thin specimen with low contrast: staining. Selective stains react differently with different parts of the specimens. Certain DNA stains (careful, potentially carcinogenic!) interact with the DNA and make nuclei visible. Other stains interact with other substances. Here it is necessary to consult a catalog to determine the right stain for the task.
• Thin specimen with low contrast: observing in phase contrast. Phase contrast microscopy is an optical method in increase contrast. A prerequisite is, that the specimen possesses a different refractive index than the surrounding medium, which is the case most of the time.
• Thick and soft specimen: squeezing. The specimen can be squeezed between the slide and the cover glass. One example of this method is the observation of various fruits, such as a soft kiwi.
• Thick and soft specimen: hardening followed by microtoming. Soft specimens (ripe fruits, soft leaves etc.) are often difficult to cut into thin sections. They have to be hardened first. Plant materials can be hardened by placing them into alcohol for a few days. This removes water and makes the object easier to cut into small slices. Be careful again, this method is not suitable for children, due to the sharp tools involved. Also note, that the removal of water by the alcohol may cause the specimen to shrink.
• Thick and hard specimen: softening. Certain specimens can be softened by boiling them. Alternatively, certain chemicals also achieve the same effect. The soft specimen can then be squeezed between the slide and cover glass before microscopic observation.
• Thick and hard specimen: grinding them thin. This method is sometimes used when observing rocks and other hard substances which can not be softened. Specialized tools are required.
• Thick and hard specimen: use stereo-microscopes. The easiest way is to observe them with a stereo microscope using epi-illumination (light from the top).

When the glue has dried completely, carefully peel off the glue. It should separate easily from the leaf. The leaf has left an impression on the glue.

Cut the glue into shape using scissors and observe it with the microscope. If the glue is still water soluble after drying, then do not immerse the glue into water. The contrast is low, it is necessary to close the condenser aperture diaphragm.

Materials: Leaf of a plant, white wood glue (PVC glue etc., water soluble), slides, scissors.

Method:

1. Evenly spread a drop of water soluble wood glue on the bottom side of a leaf (the stomata are located on the bottom side).
2. Wait several hours or overnight for the glue to dry.
3. Carefully peel off the glue. It has become transparent.
4. Use scissors to cut the glue into shape and observe under the microscope. The leaf epidermis cells have left an impression on the glue, which can be observed.

Troubleshooting:

Problem: The glue does not want to separate from the leaf
Solution: Spread the glue on an even section of the leaf underside. Some leaves may have microscopic hair, which have become attached to the glue.

Problem: Nothing can be seen.
Solution: The contrast of this specimen is very low. You have to close the condenser aperture diaphragm to increase contrast.

Problem: The resolution is low.
Solution: This is due to the fact that the specimen (the dried glue) is not embedded in water and a cover glass is missing. Either make a permanent mount in with a non water based mounting medium or try to use glue which is not water soluble anymore after it has dried.

Issues to consider:

• Do not cover the whole underside of the leaf with glue, this will block gas exchange and may harm the plant.
• Do not use glue with organic solvents (acetone, alcohols etc.). This will possibly harm the leaf and these solvents withdraw water from the cells and dissolve the cell membrane. Or: try it anyway, maybe it still works… Take care that the glue does not contain solvents that are harmful when inhaled.

Things to try (I never tried them, success not guaranteed!):

• Spread the glue at night (do not turn on the lights) and compare the shape of the stomata with those during day. The stomata of the “daytime glue” should be open, the stomata of the “night time glue” should be closed.
• Compare the size and shape of the leaf epidermis cells of different plants.
• Does the size of the leaf have an effect on the number of stomata, on their shape?
• Approximately how many epidermis cells are there to one pair of guard cells?

Wikipedia explanation of stomata and guard cells.

This picture shows the tip of a maple leaf. Note that not all leaves can be processed this way.

You may be interested in the “Virtual Microscope”, which allows you to zoom into the leaf veins: Virtual microscope: maple leaf skeleton

Materials: Maple leaves, hot plate, cooking pot, eating plates, small but stiff brush or toothbrush

Method:

1. Let the leaves simmer for 1-2 hours. Periodically check the leaves by carefully rubbing them between your fingers. They should start to feel slimy and you should be able to rub off some of the surface plant tissue.
2. Carefully lift out the leaves. They are now very delicate and they tear easily. Put one leaf on one dish each.
3. Add a bit of water to the leaf on the dish. Use the brush to carfully remove the soft plant tissue of the leaf. The brush presses the leaf against the plate. This gives the leaf stability. Use the fingers of the other hand to prevent the leaf from moving while brushing. The leaf veins start to appear. Carefully turn the leaf around and remove the plant tissue on the other side as well. The water of the dish starts to accumulate plant tissue and should be exchanged periodically.
4. You now have a delicate network of leaf veins on the plate. Lift it out and place it flat on tissue paper to remove most of the liquid. Press the leaf veins between layers of tissue paper and a book. Otherwise there is the danger that the leaf will warp during the drying process.
5. Observe the leaf veins using a stereo microscope. They can also be observed using a compound microscope using a low magnification. Alternatively it is possible to scan the leaf veins with a flat-bed scanner.
6. Make a quality check. Observe any soft leaf material that has not been removed. Observe any tears and breaks in the leaf veins that were caused by brushing too forcefully.

Alternative method:

• Press the leaf between two books.
• Place the leaved into a solution of washing soda (pH 11 – don’t let children do this!) until they become pulpy and the soft material starts to come off.
• Rinse the leaves and brush off the soft material with a soft brush.

The Efficient Method: Do an Internet search for “skeleton leaves” and buy some ready made ones…

Other Ideas:

• Students may also attempt to remove the soft tissue directly under the stereo microscope. In this case the leaf should be placed in a petri dish.
• The cleaned leaf veins can be brightened by washing them in pure alcohol. This removes remains of the chlorophyll. The alcohol also removes water and the network of veins will shrink. Wash the veins in pure water after the alcohol treatment to restore the original size.
• The network of veins can be scanned using a flatbed scanner using high resolution. This also visualizes small structures. A dark background gives a nice contrast.

Troubleshooting:

Question: It is not possible to remove the soft tissue of the leaf.
Answer: Some leaves can be boiled for hours and still not macerate. Oak leaves are completely unsuitable for this preparatory technique. Try out a variety of different leafs. Alternatively, the leaf may not have been boiled long enough.